"how long does a frozen muscle have to be in
the -70 freezer before you can cut it on the cryostat? "

**** Doesn't have to be at
-70oC for a specific period of time. Have cut and stained frozen sections immediately after freezing as well as several months at
-70oC

Do any of
you use control slides when staining? And if you do use controls do they have
to be fresh tissue or paraffin embedded tissue?

No use inbuilt controls

The
stain giving us the most problems is the ATP. We can't seem to get it to work
consistently. We also are not doing the pH, and wonder how important that is
to the staining process? I'm thinking pretty important.

The staining protocol we are using is as follows: 1. Slides
in Sodium barbital buffer for 10 minutes. 2. incubate slides in incubation
solution at 37 degrees c. for 30 minutes. 3. Transfer slides to 2%
cobaltous chloride solution for 3 minutes. 4. Wash slides in distilled
water for 5 minutes. 5. we then dip slides (one by one) in Ammonium sulfide 6-7 times, sometimes more. I believe the stain gets darker the more
you dip. 6.Rinse in water and check differentiation. At this Point I expect to see two different shades of brown. 7. Wash in running tap water
for 20 minutes, if everything looks like it should. 8.Deydrate in graded
alcohols, clear in histoclear or xylene and coverslip in synthetic mounting
medium.

The solutions are made the following way:

Sodium barbital buffer: sodium barbital 2.062g calcium chloride .999g d.
water 1000 ml The ph is then supposed to be adjusted to 9.4. Stable for 2
months. Store in refrigerator. We do not do anything with the pH because
this is not how we were taught to do the stain.

2% Cobaltous chloride:
Cobaltous chloride 2.0g d. water 100 ml

Ammonium
sulfide: Ammonium sulfide 1.0 ml tap water 10 ml (but I was taught to use 20 ml) I don't know why. Does this need to be made fresh each
time? Should the Ammonium sulfide be stored in the dark?

Incubation
Solution: Sodium barbital buffer 10.0 ml Adenosine-5-triphosphate (ATP) 15mg This also needs a pH of 9.4 and is prepared fresh each time.
Again no pH modifications are done.

Basically when I do this stain it
never looks the way it looks in the textbook. It always looks mottled. It never looks crisp with a distinct color difference.

Does this stain
react differently with different muscles? If the muscle sample is of poor quality does it affect the staining?

I am also having trouble with the
Modified Gomori Trichrome:

how important is it to stain the slides
within one hour of cutting? - I have not found it
important

does anyone filter the trichrome solution
before staining? - yes at least
once/week

I find this stain very basic yet it never
comes out consistently. As of late, there is a dark , black precipitate
like artifact on the slides after staining. I'm not sure whats causing
it. - probably Haematoxylin ppt. Try filtering the
Haematoxylin

Stain
slides in Harris Hematoxylin for 5 minutes. Rinse in Distilled water to remove excess Hemo. Stain in trichrome solution for 20 minutes.
wash in distilled water to remove excess trichrome. Dip twice in 0.2%
acetic acid. ( we sometimes use 3%). Dehydrate in graded alcohols and clear in xylene. Mount in synthetic medium.

I'll get back to everyone
on the Nadh, Pas, Pasd, oro later. I don't want to bombard everyone. Feel free
to post these on the histonet if its easier for you. Thanks to everyone who
responded. I appreciate it.

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